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Abstract

Summary

Transcription factories are nuclear domains where gene transcription takes place
although the molecular basis for their formation and maintenance are unknown. In this
study, we explored how the properties of chromatin as a polymer may contribute to the
structure of transcription factories. We found that transcriptional active chromatin
contains modifications like histone H4 acetylated at Lysine 16 (H4K16ac). Single
fibre analysis showed that this modification spans the entire body of the gene.
Furthermore, H4K16ac genes cluster in regions up to 500 Kb alternating active and
inactive chromatin. The introduction of H4K16ac in chromatin induces stiffness in the
chromatin fibre. The result of this change in flexibility is that chromatin could
behave like a multi-block copolymer with repetitions of stiff-flexible
(active-inactive chromatin) components. Copolymers with such structure self-organize
through spontaneous phase separation into microdomains. Consistent with such model
H4K16ac chromatin form foci that associates with nascent transcripts. We propose that
transcription factories are the result of the spontaneous concentration of H4K16ac
chromatin that are in proximity, mainly in cis.

Background

Transcription in eukaryotes is organized in transcription factories (TFs), which are
nuclear domains where several genes are grouped to be transcribed together [1,2][3-5]. The current opinion is that the genes in a TF interact by a looping
mechanism [5-8]. It has been suggested that chromatin looping plays an important role in
controlling gene activity by bringing together promoters and enhancers or TFs [9]. Some studies suggest that promoter-enhancer loops are maintained by the
interaction of proteins associated with these cis-regulatory elements [10]. This interaction precedes chromatin activation, which is required for gene
relocation to the TF by an unknown mechanism [11]. It has been proposed that TFs are maintained by depletion attraction forces
(excluding volume effect) between RNA pol II molecules [12]. However, experimental evidence has shown that genes remain at the factory
even when active RNA pol II is not present [13]. This makes the excluding volume model very improbable and suggests that this
structure is not the result of transcription. Instead experimental evidence points to
histone acetylation as being responsible for loop formation [11]. For these reasons we explored the possible contribution of chromatin
acetylation in the formation of TFs.

Chromatin at the TF is decondensed [2] and contains active transcription marks like histone acetylation or H3K36me3 [14]. Among all the possible Lysine residues that can be acetylated, H4K16Ac is
very special because it prevents the formation of compacted chromatin by inhibiting the
inter-fibre interaction [15-18]. Moreover, H4K16 acetylation is associated with both active chromatin [19] and with the active transcription marker H3K4me3 [19-21].

Results and discussion

To confirm whether H4K16Ac is associated with active chromatin, we analysed the
distribution of H4K16Ac in the nucleus of the TFs of circulating lymphocytes. The TFs
were visualised as sites of incorporation of Br-UTP into nascent RNA. TFs appeared as
discrete foci distributed along the edge of condensed chromatin (Figure ​(Figure1a)1a) as previously described in other cell types [2,22,23]. H4K16Ac was scattered in foci overlapping or very close to these Br-RNA
sites (Figure ​(Figure1a).1a). To study the extent and degree of the
hyper-acetylated chromatin in individual transcription units (TUs), we deconstructed the
nuclei of these cells by making chromatin spreads. Under these conditions active RNA
polymerases and epigenetic modifications of the chromatin are preserved. This treatment
disassembled nuclei and spread templates over a wide area. The DNA adopts a linear
structure with no visible nucleosomes [22] and 95% of active polymerases remain associated with the DNA [24]. When we stained the chromatin spreads with antibodies against H4K16Ac, they
showed almost continuous fluorescent tracks along the DNA fibres (Figure ​(Figure1b).1b). These H4K16Ac tracks corresponded with active chromatin, as demonstrated
by co-localisation of H4K16Ac with the nascent transcripts that were labelled either
in vivo or in vitro by using Bromo-Uridine (BrU) or BrUTP
respectively. The area covered by acetylated histones was larger than that stained by
the nascent transcripts (Figure ​(Figure1c).1c). This was to be expected
because histone acetylation extends over long stretches of genes, whilst only a few RNA
pol II molecules are ever found on a given gene [22]. Nevertheless, to demonstrate that the distribution of active RNA pol II
molecules is not an artifact of non-natural nucleotide incorporation, we carried out
chromatin spreads with cells where transcription was not labelled with BrU. Our
experiments demonstrated a similar co-localisation of H4K16Ac with P-RNA pol II
(hyper-phosphorylated Ser2) (Figure ​(Figure1c).1c).

Transcription on acetylated chromatin. (a) The nascent Br-RNA
(green) and H4 K16Ac (red) signals are closely associated. BrUTP incorporation in
human lymphocyte was carried out for 15 min and after fixation immunolabeled
together with histone H4 K16Ac...

The chromatin spreading technique allowed us to measure the length of H4K16Ac tracks.
The distribution of H4K16Ac stretches showed a lognormal distribution with average size
of ~15 Kb (Figure ​(Figure1e).1e). H4K16Ac tracks rarely appeared isolated,
instead they tended to cluster, spanning several hundreds of Kb (348 ± 90; range
235–530 Kb) (Figure ​(Figure1d).1d). The extension of the gaps between
two consecutive H4K16Ac tracks in the cluster showed a lognormal distribution with an
average distance of ~30 Kb (Figure ​(Figure1f).1f). The analysis of the
polymerases loaded onto H4K16Ac tracks showed that not all the tracks were stained with
Br-RNA or P-RNA pol II. The number of nascent transcripts or P-RNA pol II per track was
low (0.7 ± 1 transcripts/track and 0.8 ± 0.9 P-RNA pol II/track). This was in
accordance with our previous findings, suggesting that most of the TUs contain one
molecule of RNA pol II [22]. The fact that some H4K16Ac tracks of chromatin were not associated to RNA
pol II or Br-RNA could reflect a temporal discrepancy between the transcription and
acetylation processes of chromatin. Indeed, transcription by RNA pol II takes only a few
minutes [25-27] while deacetylation of active chromatin can take several hours [28], providing a molecular memory of recently-transcribed chromatin. On the other
hand, H4K16Ac tracks are not a special feature of lymphocytes as we were able to find
the same chromatin organisation in all the mammalian cell types tested including: Hela,
Epstein Barr transformed lymphocytes, human lymphocytes, primary human fibroblasts,
primary mouse fibroblasts and murine erythroleukemia cells (both differentiated and
undifferentiated).

The clusters in all the different cell types analysed were identical with respect to the
number of TUs (8 ± 2 TUs/Cluster), suggesting that co-linear active genes expressed
at the same time, in agreement with the analysis of expression data using FDCP mix cells [29]. The sliding window analysis (applying a window of 500 Kb) over the entire
genome showed that genes are active in clusters (Figure ​(Figure1g),1g), in
accordance with our chromatin spreads data. Moreover, our results are consistent with
the co-expression data after a Serial Analysis of Gene Expression where the cluster size
was <500 Kb [30]. From these data we can conclude that co-linear TUs are active at the same
time in the same cell.

How are these TUs organised in the cell nucleus?

Collinear active TUs are enriched in H4K16Ac which confers stiffness and inhibits
inter-fiber interaction [15-17]. In this way, chromatin appears as a multi-block copolymer with stiff and
flexible monomers (rod-coil)n system, where the rod is the stiff active
TU. The multi-block copolymers function as amphiphiles whose components segregate
into domains to avoid unfavourable contact with each other. In these systems,
complete phase separation is prevented by the covalent linkage between the components [31]. The rod block does not have the same conformational entropy as the coil
block and this restricts homogeneous packaging. In consequence anisotropic
interactions occur between the stiff blocks ending in a liquid crystalline domain
where the different TUs are aligned in a high order smectic phase [31] (Figure ​(Figure2a).2a). Multi-block copolymers can adopt many
different structures depending on the relative proportions of the rod and coil
phases. For example, when the rod phase is lower than 20% the structures obtained are
microspheres [32] (Figure ​(Figure2b).2b). A calculation of the amount of active
chromatin in a Hela cell line gives a proportion of rods to coils of ~12%, which is
consistent with active chromatin separated in many microspheres. These spheres for
H4K16Ac chromatin were observed in the cell nucleus of human lymphocytes (the shape
factor was 0.93 + 0.05). Microspheres are regularly distributed in artificial
polymers with regular coils and rods resulting from the repulsion forces of coils
pushing in all directions. In the cell nucleus TFs are not regularly distributed
because the sizes of genes and intergenic distances are not as regular as in
artificial polymers. Moreover, H4K16Ac foci concentrated at the edge of condensed
chromatin (Figure ​(Figure1a).1a). A possible reason for this discrepancy
is that chromatin cohabits with the inter chromatin compartment (ICC), which is
composed by RNPs and proteins. This results in a biphasic system where the ICC
(inelastic phase) segregates from chromatin [33], creating an interphase between both components. Under these conditions
microspheres containing H4K16Ac may be pushed by the coil polymers to the interphase
between chromatin and ICC (Figure ​(Figure2c),2c), resulting in the
localisation along the edge of the chromatin as observed.

A prediction of the multi-block copolymer model is that microphase separation must
persist as long as H4K16Ac is present in chromatin. In fact, H4K16Ac foci were
unperturbed by treatments like 2 M NaCl extraction, which disrupts chromatin;
transcription inhibition by DRB (5,6-Dichlorobenzimidazole
1-β-D-ribofuranoside), which reduces RNA pol II transcription by 98%; and heat
shock (1h 45°C), which releases RNA pol II from the DNA [26] (Additional file 1: Figure S1). The only way to
disrupt these foci was by formamide treatment, which works as a solvent for the
electrostatic self-assembled polymers (Additional file 2:
Figure S2).

These experiments contradict the excluding volume model [12] and are in agreement with the multi-block microphase separation hypothesis
proposed in this study.

The next question about the genes in a TF is where they come from. Several studies
have shown that genes in cis and in trans are able to interact in the same TF [5-7]. However, the analysis of chromatin spreads showed that collinear genes
are active in the same cell at the same time. This guarantees that several H4K16Ac
tracks are in close proximity. Therefore, most of these collinear TUs would probably
aggregate in the same microsphere, as occurs in similar situations with the
multi-block copolymers [34]. The experimental evidence from chromosome configuration capture analysis
suggests that local chromatin is the primary source of interaction for any genomic
loci [8]. Nevertheless, we cannot exclude the possibility that some genes located
further away in the same chromosome or in another chromosome can interact due to
proximity or chromatin folding.

Finally, a remarkable feature of TFs is their constant size across species and
differentiation stages [23]. According to the multi-block copolymer model for chromatin organisation,
the way to change the size of H4K16Ac foci (and consequently TFs) is by increasing
the number of active genes in a given region or by unrestricting the mobility of the
active chromatin. The latter has been reported in experiments using plasmids that
rendered larger TFs than the endogenous ones [35,36].

In summary, we present evidence of the relationship between epigenetic marks and the
TF structure. Our model proposes that active chromatin self-organises in the nucleus
due to the special physical properties of H4K16Ac modified chromatin. Therefore, our
model implies that chromatin becomes activated (H4K16Ac modified) before joining a
TF. This is conceptually very different from current transcription factory model,
which proposes that genes are targeted to TFs to “enhance production by
concentrating the relevant machines, resources, and expertise in one place” [37].

Materials and methods

Transcription in vivo and in vitro

For in vivo transcription, cells grown on coverslips were incubated in presence of
2.5 mM BrU for several min.

For in vitro transcription, cells grown on coverslips were treated as described [3].

Chromatin spreading

Cells (103 cells in 5 μl) were spotted onto a 22 × 50
mm glass slide and 5 μl of lyses buffer were added (Lyses buffer: 1%
sarkosyl, 25 U/ml ribonuclease inhibitor, 10 mM EDTA, and 100 mM Tris–HCl (pH
7.4)). After 10 min at 20°C, the slide was tilted to allow the drop to run down.
Samples were air-dried and fixed in 4% Paraformaldehyde for 10 min. Clusters were
defined as two o more hyper acetylated tracks in less than 100 Kb. For quantification
of clusters of hyper acetylated chromatin between 150 and 200 tracks of hyper
acetylated chromatin were analysed.

The degree of spreading of the chromatin was measured by hybridising the spreads with
a fragment of DNA of 47.26 Kb; the spreading was 3.9 + 0.2 Kb/μm.

Microarrays and sliding window analysis

Mouse FCDP mix cells were used. cRNA synthesis and hybridisation to oligonucleotide
array were performed as described [29].

The sliding window analysis was performed by applying a window of 500 Kb over the
chromosomes and moved at 5 Kb steps along a chromosome to know whether the genes
contained in that window were more likely to be transcribed together than just by
chance.

Authors' contributions

AZC-H, RPN and JER performed some of the immunocytochemical experiments and acquired
data. SS and TE, performed the transcriptomic and the statistical analyses. CI
participated in the draft the manuscript. VJB was in charge of the DNA fish experiments
& FJI conceived of the study, and participated in its design and coordination and
helped to draft the manuscript. All authors read and approved the final manuscript.

Supplementary Material

Additional file 1: Figure S1:

Stability of H4 K16Ac foci. Resistance of H4 K16Ac foci to various treatments
that disrupted transcription or chromatin structure. The aspect of H4 K16Ac
foci did not change after DRB treatment (2h 150 μM) or heat shock
(Hs) for 1h at 45°C. Both treatments led to the release of RNA pol II from
the genes. These foci were also resistant to NaCl extraction (cells
permeabilised with 0.05% Triton X100 for 5 min in PBS at 4°C followed by
10 min extraction with 2M NaCl for 10 min). The images were pseudo-coloured for
display. The bottom bar shows the scale of pseudo-colours used.

H4 K16Ac foci are disassembled by formamide. The resistance of H4 K16Ac foci to
formamide treatment. Cells were incubated for 5 min in PBS with different
concentrations of formamide (0, 25, 50 and 100%) then fixed with 4%
paraformaldehyde and immunolabelled with H4K16Ac antibodies. (a) The
H4K16Ac foci were disassembled by formamide treatment, as can be seen from the
change in the staining pattern, which is more diffuse and less intense than the
control. The images were pseudo-coloured for display. (b) The
deconstruction of the foci was quantified by the change in the pixel intensity
variation coefficient (SD/mean). This analysis was performed by measuring the
mean intensity and the standard deviation (SD) of the H4K16Ac signal of the
nuclear areas in at least 200 cells for each treatment. The images were
pseudo-coloured for display. The bottom bar shows the scale of pseudo-colours
used.

Acknowledgements

The authors would like to thank Elspeth McFarlane for helping with the English. To:
Ministerio de Economia y Competitividad, (Spain) (Grant number: BFU2009-10792) and
The Medical Research Council (UK), for supporting this work. We also thank
Fundação Ciência e Tecnologia, Portugal for funding RPN.